The pretreatment method in marine organisms and sediment for microplastics analysis by FTIR using “Cylindrical microplastics fractionator”

For the detection of microplastics (MPs) in aquatic biota using Fourier transform infrared spectroscopy (FTIR), the ability to remove organic matter (OM) in pretreatment steps is essential to increase the time efficiency of MPs measurement and method uniformity. In principle, decreasing OM can be achieved by increasing the number of pretreatment steps. However, MPs are lost in proportion to the number of transfers between each step. Therefore, we have created a "Cylindrical MPs Fractionator" composed of commercially available materials. This container allows for a six-step pretreatment process that is designed to increase the removal capacity of OM with only one transfer to prevent the loss of MPs.• Biological or sediment samples are placed in the extractor and subjected to chemical treatment and density separation.• Residues containing MPs are obtained on filters by vacuum filtration.• After additional chemical treatment of the obtained residue, the components of the residue are identified by microscopic FTIR.This method removed 99.3% of OM and recovered 88.5% of MPs. The presenting method confirmed that this can be used with the same process for 11 organisms and sediments from estuarine ecosystem in Japan as models.

• Biological or sediment samples are placed in the extractor and subjected to chemical treatment and density separation.• Residues containing MPs are obtained on filters by vacuum filtration.
• After additional chemical treatment of the obtained residue, the components of the residue are identified by microscopic FTIR.
This method removed 99.3% of OM and recovered 88.5% of MPs.The presenting method confirmed that this can be used with the same process for 11 organisms and sediments from estuarine ecosystem in Japan as models.

Specifications Table
Subject area: Environmental Science More specific subject area: Microplastics analysis Name of your method: MPs pretreatment method for aquatic organisms and sediment Name and reference of original method: NA. Resource availability: Reagents and Equipment are mentioned in the article.

Overview
The pretreatment method for microplastics (MPs) analysis using the "Cylindrical MPs fractionator" is presented here.This method assumes the biological sample consists of four components: proteins, carbohydrates, fats, and carbonate minerals.First, carbonate minerals are removed by formic acid.Then, the remaining three components are decomposed by Fenton's reagent.However, since the specific gravity of the lipids is low compared to the specific gravity of Fenton's reagent, the components tend to be residual at the inner wall during the reaction.This is caused by the oxygen bubbles' upward action pushing them up the sample.Therefore, potassium hydroxide is used to saponify the remaining unreacted adipose tissue.Add Ethylenediamine-N,N,N',N'-tetraacetic Acid Disodium Salt Dihydrate-2Na (EDTA-2Na) simultaneously with potassium hydroxide to dissolve the carbonate that could not be removed with formic acid.This causes most of the biogenic material to be micronized and its breakdown products, together with iron oxide (or iron hydroxide), to form a sludge.The sludge is then settled by density separation and is removed.However, during the density separation process and filtration, some organic matter (OM) remains on the filter.Particularly, fibrous filters collect degradation products smaller than the pore size.This can result in a coating on the microplastic that prevents the infrared absorption spectrum of pure microplastic from being obtained.To further remove residues, the filter is rinsed with a formic acid solution during decompression filtration (additional demineralization), lipids and proteins are emulsified with sodium dodecyl sulfate (SDS), and polysaccharides are removed with a NaOH/Urea/Thiourea (NaUT) solution ( Table 1 ).Finally, after rinsing with distilled water, the filter is fixed between two glass slides with a silicone O-ring.This prevents the contamination and contraction of the sample during drying ( Fig. 1 ).After thoroughly drying, the sample is subjected to Fourier transform infrared spectroscopy (FTIR) analysis.

Overview of the fractionator
The fractionator is designed to meet two specific conditions.First, the material must be highly resistant to various chemical treatments, heating, and density separations.Second, the internal structure must be cylindrical to not interfere with the particles' vertical movement.The overall view of the fractionator is shown in Fig. 2 (a) and (b).The pipe is made of perfluoro alkoxy alkane (PFA) resin ( Fig. 2 c).The upper plug is usually an aluminum cap ( Fig. 2 d).The lower plug has a hole drilled with a diameter of 18 mm ( Fig. 2 e).During the pretreatment process, the sample is first put into the lower plug.It is used as a receptacle to measure the mass of the sample; the plug is then inserted into the pipe.During the density separation step, the upper plug with a valve is used instead of the aluminum cap ( Fig. 2 f).Adjust this plug-mounted valve to release the lower part of the solution from which the sludge has settled.Air flows into the pipe through a syringe filter, so no contamination of MPs occurs.The fractionator can be substituted by turning a cylindrical separatory funnel upside down.However, the joint of the upper stopper (lower stopper when in use) of the separatory funnel should be fitted with a PFA sheet to prevent adhesion by the alkaline solution.Using the separatory funnel in its normal orientation is not recommended because sand may clog the cock during density separation.

Materials and budget of the fractionator
The original materials for this fractionator can be purchased mainly from Flonchemical Co., Ltd.A set of fractionators costs approximately US$165 (US$1 = ¥150, not including the cost of tools needed to process the materials).The fractionator can be cleaned and reused like regular laboratory equipment.Considering that the cock valve is used only for density separation and, therefore, not required for every fractionator, the cost for the second and subsequent sets will be less.Prices of materials are constantly changing and should be checked accordingly.Note that materials need to be purchased in unit lengths.

Experimental preparation and precautions
Experimenters should wear clothing that does not contain plastic fibers (e.g., 100% cotton) whenever possible and white lab coats with minimal skin exposure (safety considerations).Appropriate protective equipment (e.g., gloves, goggles) should be used when using hazardous reagents.Clothing should be carefully cleaned of foreign particles using adhesive tape prior to MPs extraction procedures.
The following steps are required before extraction: 1) Preparation of laboratory equipment: All laboratory equipment must be washed/rinsed with distilled water immediately before use.All extraction procedures must be performed under sterile and clean conditions (using of a dust-proof clean bench is recommended).2) Record the mass of the pipe (-> A), aluminum cap (-> B), and lower plug (-> C) (these will be needed to calculate the volume of solution to be added during density separation).3) Fill the hole in the lower plug with distilled water, transfer this to a graduated cylinder, and record the hole volume (-> D).Fig. 3 shows the whole process abstractly.The processing procedures for biological and sediment samples are identical.An experimental log sheet is depicted in supplemental materials.

Carbonate minerals elimination [Step 1 in Fig. 3 ]
Formic acid is used for carbonate minerals elimination.Invertebrates may have a complex junction of exoskeleton and viscera, and removal of the exoskeleton by dissection prior to pretreatment may lead to loss of viscera containing MPs.Therefore, it may be better not to remove the shell and skeleton before pretreatment.When the pretreatment is conducted with the exoskeleton, the formic acid will first remove the carbonate from the exoskeleton, allowing subsequent solutions to penetrate more easily into the interior.In the case of crustaceans, they contain calcium carbonate as part of their chitin.Carbonate minerals elimination at the beginning of the process will enhance the ability of the chitin to break down.

1) Preparation of carbonate minerals elimination reagent:
There are two carbonate minerals elimination regents,  and . is used before the hydrogen peroxide treatment, and  is used in the filtration.The prepared solution is filtered through a filter syringe (or reduced-pressure filtration) with a smaller mesh than the filter used in MPs collection.
• 3-Methyl-1-butanol ( "3M1B ", 40 μL / mL CAS RN®: 7782-63-0) * This is also an antifoaming agent and is added to allow hydrogen peroxide to react safely.However, D5 and PG have long-lasting antifoaming effects, and they have the property of temporarily increasing the amount of foam.To reduce this, 3M1B is added to promote defoaming due to its Marangoni effect.Depending on the experimental conditions, the defoaming effect of 3M1B does not last for several days.Therefore, it is recommended that it be used in combination with D5 or PG.Ethanol can be substituted, although it has a shorter lifespan.Test it once with adequate safety precautions, check the foam generation, and adjust the amount of defoaming agent and the temperature increase program (Step 2).This mixing should be done immediately before use.
2) Place the sample in the lower stopper and measure the mass.* Perform dissection as appropriate.Refer to Table 3 to determine if dissection is necessary.
3) Insert the lower plug into the pipe body.4) Add 3 mL of carbonate minerals elimination reagent  per 1 g of the sample and put the upper cap on.5) Wait until the bubbling is stopped.* The end of bubbling may not be due to the end of carbonate minerals elimination but rather to the complete decomposition of formic acid.Therefore, add a small amount of additional formic acid solution to confirm that carbonate minerals elimination has been completed.6) When the carbonate minerals elimination is finished, add another 0.1 mL of carbonate minerals elimination reagent  and go to the next step.Organic decomposition is performed using Fenton's reagent.Immediately after adding the reagent, the reaction may heat up, so ice-cooling is performed.Once the reaction cools down, heat to 50 °C to accelerate the reaction.At this time, the fractionator is immersed in water to prevent it from exceeding 50 °C due to reaction heat. 1) Prepare ice water in a 1-2 L beaker.
2) Add 30 mL of 35% hydrogen peroxide solution (CAS RN®: 7722-84-1) per 1 g of sample.* Add in small quantities to avoid strong reactions.Do not approach the body or look inside from above.
3) Immerse the fractionator in ice water and allow it to stand for 24 hrs.( Fig. 4 , The ice will gradually thaw but should not be replaced.)4) Agitate with a mixer.* Agitation is not mandatory, but it allows the tissues to mix uniformly with the solution, thus promoting an efficient reaction.However, be careful that the sample does not adhere to the vessel wall after stirring.If the sample adheres to the wall, open the cap on a clean bench and drop the sample into the solution using a long-tipped metal spatula.The adhesion of organic matter to the wall surface that occurs at this time will greatly affect the efficiency and accuracy of the subsequent FTIR analysis.5) Allow to stand for 24 hrs.at room temperature while immersed in room temperature water.6) Allow the external container containing the water to stand at 50 °C for 72 hrs.7) Bring to room temperature and go to the next step.

KOH process [Step 3 in Fig. 3 ]
Organic decomposition is performed using alkaline solutions.The main purpose is to decompose diatoms and saponify lipids.If the hydrogen peroxide is still active in Step 3, the reaction proceeds rapidly, so add hydrogen peroxide in small amounts while monitoring the condition of the sample.If the reaction is too strong, stop adding hydrogen peroxide and allow it to stand for several days until the hydrogen peroxide is gone.
3) Incubate at 50 °C for 1 day.4) Bring to room temperature and agitate the sample with a vortex mixer or the like.5) Go to the next step.Density separation is performed using a sodium iodide solution.Mineral particles in the sample are removed in this step.The specific gravity of the internal solution is first calculated, and the final density in the solution is adjusted to 1.6 by adding a sodium iodide solution with a density of 1.8.When placing the fractionator, stand it vertically to not interfere with the vertical movement of the particles.The operation is repeated three times.
* Caution!Mixing sodium iodide and hydrogen peroxide causes an explosive reaction.Use the highest possible precautions to avoid mixing the two.
1) Prepare sodium iodide solutions with specific gravities of 1.8 and 1.6.Place a magnetic stirrer in a 1 L beaker and add 500 mL of distilled water.Add sodium iodide (CAS RN®: 7681-82-5) and stir until dissolved.Using a graduated measuring cylinder, measure 10 mL of the solution to determine the specific gravity.Continue until the solution reaches the desired gravity.* A minimum of 1 L of sodium iodide solution with a specific gravity of 1.6 is required.
2) Filter the solution through a filter paper with a pore size smaller than the smallest particle size of the target MPs.
3) Measure the weight of the fractionator with the sample.-> E 4) Measure the height of the sample solution.-> h 5) Estimate the specific gravity of the solution using the following formula (Refer to the pretreatment log table as necessary): where the  is the pipe's inner diameter.
6) Add 1.8 sodium iodide so that the final specific gravity becomes 1.6 by the specific gravity of the solution obtained by the following formula: 1 .8  ℎ   = 5 ( 1 .6 −  )  7) Add 1.6 sodium iodide to bring the volume to 150 mL.8) Stand the fractionator upright and allow it to stand for at least 3 hrs.9) Fill a 1 L beaker to the 600 mL with a sodium iodide solution of specific gravity 1.6.10) Loosen the stopper at the bottom of the fractionator to the extent that the container does not leak, and close the valve at the top.11) Immerse the lower part of the fractionator in 1.6 sodium iodide and gently open the loosened lower stopper in the solution.12) Open the upper valve slightly, pour half of the sodium iodide in the fractionator into the external solution, and close the valve again.13) Close the lower part with another clean plug and open the upper valve.14) [First and second times] Add sodium iodide solution with a specific gravity of 1.6 to 150 mL.15) [Third time] Go to the Filtration without adding sodium iodide solution.

Filtration
Filtration is performed using a stainless-steel mesh and a PTFE filter ( Fig. 5 ).Use a funnel for the stainless-steel mesh and a vacuum filtration device for the PTFE filter.If the PTFE filter becomes clogged, carbonate minerals elimination can be induced again with a formic acid solution, and filtration can be restarted or moved to the next step.
* There are two reasons to use more than one filter type.One is the infrared absorption spectrum.Thick samples may not transmit infrared light, and larger particles are measured by the reflection method.The second is a filter area issue.In our protocol, we set a PTFE filter in a filter holder with a funnel diameter of 4 mm for filtration to reduce the FTIR measurement time as much as possible.The presence of large MPs, even a small number, may cause sample overlap and interfere with the detection of small MPs.For this reason, the filtration in this study follows this step.This is not mandatory and can be modified to suit the experimenter's desired particle size range of MPs.
3) Co-wash the fractionator and lower stopper with distilled water.The infrared absorption spectrum of lipid content is similar to polymers, and residual lipid content can greatly interfere with microplastic measurements.The light-specific gravity of the lipid can cause it to be lifted by the bubbles of hydrogen peroxide water.Thus, SDS solution is added to emulsify the lipid content. 1) Take 1 mL of 1% SDS solution with a filter syringe and add it to the filter holder.Cover the filter holder with aluminum foil or the like and allow it to stand for 2 hrs.under 50 °C conditions.2) After 2 hrs., attach the filter holder to a vacuum filtration bottle and perform vacuum filtration again.* If lipid content were present, the filtration speed would be faster than before treatment.
3) Wash with distilled water and go to the next step.

Carbohydrate process [Step 6 in Fig. 3 ]
This treatment is intended to specifically remove chemically stable natural polymers.It is effective for samples with high woody debris content, such as riverine debris.As well as samples with robust chitin skeletons [1] .
1) Pre-cool the sealed container supporting the filter holder to − 20 °C beforehand.
3) Take 1 mL of NaUT solution with a filter syringe and add it to the funnel.Cover with aluminum foil or the like and replace the filter holder with a pre-cooled airtight container, freeze at − 20 °C.4) When the solution solidifies, remove the filter holder, and stir with a vortex mixer or the like, grasp the filter holder firmly by hand to prevent it from shifting.5) When the solution dissolves and returns to liquid, filter it under reduced pressure.6) Wash with distilled water.7) Remove the PTFE filter from the funnel and store it.* PTFE filters curl up when dried, so the edges must be restrained.They should also be sealed to prevent contamination and sa mple loss (see Fig. 1 ).

LOD, LOQ, and target polymer type of MPs
We performed a blank test ( n = 8) in our experimental environment.As a result, we observed three polystyrene particles from one blank.Therefore, we used the mean + 3 (standard deviation) and mean + 10 (standard deviation) used in general chemical analysis to determine that the limit of detection (LOD) = 4 and the limit of quantification (LOQ) = 11 particles.This varies with each researcher and experimental environment, so it is recommended to check this on a case-by-case basis.
The polymer component of interest is determined primarily based on the specific gravity limitation in the density separation.This method cannot detect fluoropolymers and epoxy resins, which may have specific gravities greater than 1.6.It also cannot detect some melamine, phenolic, and silicone resins, which have a wide range of specific gravities [2] .Furthermore, because silicone resins were used in the experiment, contamination of silicone resins could occur in principle.However, this was not observed in our blank experiments.

Recovery ratio of MPs
Spike and recovery tests were conducted with spherical polyethylene particles of different particle sizes (CPMS series, Cospheric LLC) to evaluate the recoverability of plastic particles through this method, and this was done according to the recommendations of Way et al., (2022).45-53 μm (30 particles/test), 90-106 μm (20 particles/test), 150-160 μm (20 particles/test), 180-212 μm (20 particles/test), 300-355 μm (15 particles/test), and 500-600 μm (10 particles/test) particles were treated (section 2.2).The recovery (%) was recorded from the known number of particles added using the number of particles on the filter.The tests were carried out randomly by five experimenters.For the relationship between particle size and recovery, we created a regression curve for an asymptotic regression curve through the origin.This is shown in the following equation: where Asym indicates the horizontal asymptote, R indicates the intercept, e indicates Napier's constant, and the natural logarithm of C indicates the rate constant.We received the following results ( Table 2 and Fig. 6 ).
According to Way et al. [3] , The average recovery of MPs smaller than 1 mm for the previous MPs analysis methods is 84.5%.This suggests that the recovery rate of this method was at the same level as previous methods.Thus, our MPs results are thought to have the same levels of data quality as previous knowledge.Our results showed a similar trend of lower recoveries for smaller particles, as observed by other research [ 4 , 5 ].This suggests that small particles may be relatively underestimated across the study cases and emphasizes the importance of calibrating the recoverability by each particle size.

MPs destruction in pretreatment
We evaluated MPs destruction using shape indices difference between before and after treatment.Polyvinyl chloride (PVC), Polyamide (PA), and Polymethyl methacrylate (AC) particles were used.These are chosen because of the tendency of low chemical stability.Commercially available plastic boards (PVC and AC: Hikari Co., Ltd., PA: KOKUGO Co., Ltd.) were sawed and sieved to create particles (726 ± 16 μm).The evaluation used these because they are generally polymers with low drug resistance.80 particles of each plastic type were treated.After that, 50 particles of each were randomly selected and evaluated following the procedure.Microscopic images (5440 dpi) of particles were taken using a binocular stereomicroscope (SZX10, Olympus, Japan) coupled with a camera (DP73, Olympus, Japan).Image analysis was performed using ImageJ (version 1.53a; Java1.8.0_112).The perimeter length of the particle L p , the perimeter length of the approximate ellipse L e , the major axis diameter Φ l (μm), and the minor axis diameter Φ s (μm) of the approximate ellipse were measured.The particle diameter Φ, macroscopic shape index  (0 <  ≤ 1), and microscopic shape indices  (0 <  ≤ 1) of particles before and after processing were calculated [6] : We collected the following particle shape changes ( Fig. 7 ).The results of the particle shape evaluation showed no significant changes by treatment for all three parameters except for  of AC ( Fig. 7 c). of AC showed a difference at the 5% significance level.Since no significant overall changes were observed, we conclude that the MPs are unlikely to be destroyed and are of sufficient quality for detection.However, the surface structure may be affected, so caution should be exercised in using this method when it is to be investigated in detail.

Removal ability of OM
The ability to remove OM was evaluated using two organic materials (flathead gray mullet Mugil cephalus [7] as an animal sample and common reed Phragmites communis [8] as a plant sample) to quantify the removal rate of lipids, proteins, and carbohydrates.Each test sample was ground in a mortar before being used.The lipid removal efficiency was evaluated as the palmitic acid reduction rate of M. cephalus using the gas chromatography analysis's peak area [9] .The protein removal efficiency was evaluated as the rate of total amino acids reduction of M. cephalus using an absorbance method using ninhydrin coloration [10] .The carbohydrate removal efficiency was evaluated as the rate of cellulose reduction of P. communis.This was measured by the absorbance method using phenol-sulfuric acid method coloration.Details on each of these methods can be found in the supplementary material.

Table 3
List of organisms and parts used.Second, the collection range of residue does not exactly equal the filter holder funnel shape.Due to the minute irregularities of the glass surface and the thickness of the filter itself, the actual collection area will be irregularly shaped.It is important to carefully observe the visible light image and add all particles to the FTIR measurement range, considering the possibility that MPs are collected beyond the circle range.

Limitation of sample volume
This method does not remove the solution during processing.Therefore, the disadvantage is that the amount of solution is largely relative to the amount of sample and gradually adds up.In the case of our fractionator, the maximum amount of sample that can be processed is approximately 1 g.If a sample larger than 1 g is to be processed, the only way is to separate the sample into multiple fractionators.Removal of water by freeze-drying and the creation of a larger fractionator are a couple of possible solutions.However, these have not been tried at this stage and are issues to be addressed in the future.

Fig. 1 .
Fig. 1.An overview of how the filter samples prepared are stored.The filter is sandwiched inside two glass slides (a) along with a silicone O-ring (b) by masking tape (c) or aluminum tape.The funnel size of the filter holder is 4 mm.(d) shows the assembled conservation glass slide and (e) shows the actual conservation.When the filter is larger, a larger glass slide (f) is used.

Fig. 2 .
Fig. 2. Photos of MPs fractionator.(a) The whole image of the fractionator during the chemical treatment and the static state of density separation.(b) The whole image of the fractionator during density separation.(c) Pipe body.(d) Upper cap (Inner diameter = 30 mm).To keep the internal pressure constant, the inner diameter is larger than the pipe's outer diameter; it is not sealed.(e) Lower plug.The outer diameter is equal to the pipe's inner diameter, and it is sealed.(f) Upper plug with a valve used during density separation.Air introduced from the outside passes through the filter provided and prevents airborne MPs contamination from the outside.

Fig. 5 .
Fig. 5. Images of each detailed operation in filtration step.The numbers in parentheses correspond to each filtration operation.DW: distilled water.

Fig. 9 .
Fig. 9. Photograph of a stainless-steel mesh residue sample collected from a 1 g sediment.(a): Overall view.(b): white particles on top of the black mineral particle.(c): White particles determined to be polyethylene from the same sample.

Table 1
List of each pretreatment step.

Table 2
Results of spike and recovery tests.